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Morphogenesis of the human preimplantation embryo:bringing mechanics to the clinics

Julie Firmin, Jean-Léon Maître

To cite this version:Julie Firmin, Jean-Léon Maître. Morphogenesis of the human preimplantation embryo: bringingmechanics to the clinics. 2021. �hal-03284498�

Morphogenesis of the human preimplantation embryo: bringing 1

mechanics to the clinics 2 3 Julie Firmin and Jean-Léon Maître 4 5 Institut Curie, PSL Research University, Sorbonne Universite, CNRS UMR3215, INSERM U934, Paris, 6 France. 7 Correspondence to jean-leon.maitre@curie.fr 8 9 10 11 Abstract 12 During preimplantation development, the human embryo forms the blastocyst, the structure 13 enabling uterine implantation. The blastocyst consists of an epithelial envelope, the 14 trophectoderm, encompassing a fluid-filled lumen, the blastocoel, and a cluster of pluripotent 15 stem cells, the inner cell mass. This specific architecture is crucial for the implantation and 16 further development of the human embryo. Furthermore, the morphology of the human embryo 17 is a prime determinant for clinicians to assess the implantation potential of in vitro fertilized 18 human embryos, which constitutes a key aspect of assisted reproduction technology. 19 Therefore, it is crucial to understand how the human embryo builds the blastocyst. As any 20 material, the human embryo changes shape under the action of forces. Here, we review recent 21 advances in our understanding of the mechanical forces shaping the blastocyst. We discuss 22 the cellular processes responsible for generating morphogenetic forces that were studied 23 mostly in the mouse and review the literature on human embryos to see which of them may be 24 conserved. Based on the specific morphological defects commonly observed in clinics during 25 human preimplantation development, we discuss how mechanical forces and their underlying 26 cellular processes may be affected. Together, we propose that bringing tissue mechanics to 27 the clinics will advance our understanding of human preimplantation development, as well as 28 our ability to help infertile couples to have babies. 29

Introduction: 30 In the first days after fertilization, the human embryo needs to acquire implantation 31

competencies [1–4]. The structure enabling the human embryo to implant is called the 32 blastocyst, which has a very characteristic architecture [5,6]. It consists of an epithelium, the 33 trophectoderm (TE), enveloping a fluid-filled lumen, the blastocoel, and a cluster of pluripotent 34 stem cells, the inner-cell mass (ICM). The TE invades the maternal uterus and implants the 35 embryonic tissues deriving from the ICM [7]. Until implantation, the human embryo can develop 36 without intervention from the mother. As a consequence, the self-organisation of the human 37 embryo into the blastocyst can be studied ex vivo and offers unique opportunities to study 38 human embryonic development. 39

To form the blastocyst, the human embryo initially undergoes cleavage divisions, i.e. 40 there is no cell growth during the interphase (Fig 1). Essentially, cells halve their volume with 41 each cleavage and cells rearrange themselves to shape the blastocyst without changing the 42 total cellular volume of the embryo, as measured accurately in the mouse [8]. The series of 43 cellular rearrangements sculpting the blastocyst can be broken down into three steps: 44 compaction, internalisation and lumen formation [5]. Compaction is the process by which the 45 loosely attached blastomeres enlarge their cell-cell contacts and reduce their surface exposed 46 to the outside medium [9–12]. Internalisation occurs when a subset of cells become entirely 47 surrounded by neighbouring blastomeres and isolated from the outside medium [13–17]. This 48 differential positioning of cells is a prerequisite to the first lineage specification into ICM for 49 inner cells and TE for those remaining at the embryo surface [18–21]. Finally, the blastocoel 50 appears when TE cells pump fluid, which pushes the ICM into one quadrant of the blastocyst 51 [22–24]. The lumen breaks the radial symmetry of the embryo, which is key to the formation of 52 the axes of symmetry after implantation [23,25,26]. The lumen also serves as a new interface 53 for the differentiation of the ICM into primitive endoderm (PrE) and epiblast (Epi) and of the TE 54 into mural and polar TE (mTE and pTE respectively) [27–30]. Sandwiched between the pTE 55 and PrE, the Epi will provide all cells of the human body while the PrE contributes to most of 56 the yolk sac [31,32]. The pTE is thought to mediate uterine implantation in human whereas 57 this role seems devoted to the mTE in mice [28,29,31,33,34]. 58

Since the advent of in vitro culture, the morphology of the human embryo has been 59 observed and described in numerous studies. In fact, together with cell number, the 60 morphology of the human embryo is one of the prime determinant in clinicians assessment of 61 the implantation potential of human embryos [35,36]. For example, poor compaction or slow 62 lumen growth are associated with lower implantation rates [10,37–39]. Therefore, the current 63 and future efforts in developing algorithms able to better predict the implantation potential of 64 human embryos grown in vitro will undoubtedly rely on morphological criteria [40–42]. 65

Despite the importance of human embryo morphology for its development, we know 66 little of the mechanisms responsible for shaping the human embryo [1–3]. When it comes to 67 shaping any materials, forces are necessarily in action [43,44]. In the past decades, major 68 discoveries regarding the nature of forces that shape animals were made using model 69 organisms such as the worm, fly, fish, frog and, more recently, mouse [11,45–48]. The same 70 forces are most likely responsible for the shaping of the human embryo. Of particular relevance 71 to the shaping of the human blastocyst are surface tension [49–51], adhesive coupling 72 between cells [52–54] and, osmotic pressure [55,56]. In animal cells, surface tension is 73 governed by the contractility of the acto-myosin cortex, a thin layer of cross-linked actin 74 filaments underneath the plasma membrane that is put under tension by the action of non-75

muscle myosin II motors [57,58]. The mechanical stresses exerted by the acto-myosin cortex 76 is a prime determinant of cell shape as it drives cell rounding and cytokinetic cleavage in half 77 during cytokinesis, or cell body retraction during migration or blebbing [59–63]. Adhesive 78 coupling between cells resists their separation [23,64,65]. It is mediated by cadherin adhesion 79 molecules and associated proteins, the catenins, which anchor the adhesion complex to the 80 actin cytoskeleton. By doing so, the adhesion complex connects the acto-myosin cortex of 81 contacting cells and transmits its tension throughout the tissue [66,67]. Osmotic pressure 82 dictates the movement of water between sealed compartments such as the outside medium, 83 the cytoplasm and fluid-filled lumens. Osmotic pressure relies on tight junctions, which seal 84 intercellular spaces, and ion pumps, which actively change the osmolarity of sealed 85 compartments [68,69]. Osmotic gradients will then trigger the passive movement of water 86 between compartments through aquaporin water channels [70]. 87 In this review, we discuss how learning about the mechanics of tissues can help 88 understanding the shaping of the human blastocyst. We consider how defective morphologies 89 could be explained by aberrant force patterns and point to specific cellular process underlying 90 them. In particular, we examine defects in cleavage divisions, compaction, internalisation and 91 lumen formation. Since defects in the morphology of human embryos is a prime determinant 92 of their health, bringing mechanics to the clinics will be key to improve assisted reproduction 93 technologies. For reviews covering the patterning of blastocyst lineages or general clinical 94 aspects, we recommend alternative reviews [1–4]. 95 96 Cleavages 97 Before human morphogenesis begins, cleavage divisions provide the only changes in 98 the morphology of the embryo (Fig 1). The first division takes place after a day and the second 99 half a day later [71,72]. Then, sister blastomeres divide every day in waves of progressively 100 decreasing synchrony (Fig 1). The early cleavages are notoriously error prone [4,73]. They 101 frequently lead to chromosome segregation errors [74] and are at the origin of aneuploidy that 102 can lead to developmental arrest [73]. Alternatively, these errors can be corrected by excluding 103 aneuploid cells from the embryonic tissue, which are later found in the placenta [75]. Aneuploid 104 cells can also contribute to the epiblast with or without further consequences on embryonic 105 development [76]. In addition to chromosome mis-segregation, cytokinesis can also be faulty. 106 Cytokinesis should split cells in two equal volumes, forming embryos with even cell number 107 and size. However, this is frequently not the case and has dire consequences for the embryo. 108 Indeed, human embryos with unequally sized blastomeres at the 4- and 8-cell stages were 109 found to have more frequent multinucleation and ploidy issues, which eventually reduces their 110 implantation rate [77–79]. Therefore, unequally sized blastomeres can be used as an indicator 111 of the genome integrity of the embryo and of its health in general. 112

Unequal blastomere sizes can result from distinct events (Fig 2). Cytokinesis itself can 113 produce sister cells with uneven sizes. Skewing the cleavage furrow during cytokinesis, for 114 example by contractility unbalance, will result in uneven cleavage [62]. Alternatively, 115 blastomeres can undergo “reverse cleavages”, fusing sister cells back together after the 116 cleavage furrow completed the separation of the cellular volumes of the sister cells [72,80]. In 117 addition to reducing the total expected number of cell, this results in the formation of polyploid 118 blastomeres with twice the size of their neighbouring cells, which can easily be detected. 119 Incomplete cytokinesis could result from defects in abscission, the process sealing apart the 120 cytoplasm of sister cells and leading to the formation of the midbody [81]. Abscission requires 121

the coordination of complex cellular machineries and often fails when chromosomes are mis-122 segregated. Often referred to as “direct cleavage”, blastomeres can cleave into 3 similarly 123 sized daughter cells instead of 2. After the first and second cleavages, embryos can show 3 124 cells and 5 cells respectively [72,82]. This increases the number of expected cells and results 125 in the formation of blastomeres that are 1/3rd smaller than expected, which can be difficult to 126 notice with insufficient resolution. Direct cleavages are associated with fewer embryos 127 reaching blastocyst stage and reduced implantation rate [82]. Nevertheless, blastocysts 128 produced after direct cleavages can show normal ploidy, suggesting some repairing 129 mechanism, possibly involving the exclusion of defective cells [72,83]. Such abnormal divisions 130 can result from multipolar spindle and supernumerary centrioles caused by defective sperms 131 [84,85]. Finally, human blastomeres often produce cellular fragments during cytokinesis, which 132 wastes cellular material [73,86]. Importantly, fragments sometimes entrap chromosomes, 133 making their cell of origin aneuploid [73,87]. Unsurprisingly, fragmentation is associated with 134 poor prognosis for implantation [88,89]. Fragmentation is observed frequently in human 135 embryos but rarely in mice, which makes the process difficult to study. Why human 136 blastomeres produce so many fragments during cytokinesis is unclear. Blocking the actin and 137 microtubule cytoskeletons reduces fragmentation, suggesting that these generate the forces 138 driving fragmentation [90]. Cellular fragments have been compared to blebs due to their 139 spherical shape. However, blebs are short-lived (around a minute), while fragment are stable 140 over tens of hours [60]. Blebs could persists for longer time if contractility would be hyper-141 activated [91]. However, there is not enough information available on fragment dynamics to 142 understand how they form and persist. Alternatively, membrane threads connecting the zona 143 pellucida to blastomeres were proposed to pull on blastomeres during cytokinesis [92]. 144 145 Compaction 146 Starting as early as during the 8-cell stage, compaction is the first morphogenetic 147 process associated with the formation of the blastocyst (Fig 1). During this process, 148 blastomeres get closer together, forming a tighter structure (Fig 3). This developmentally 149 regulated adhesion process requires the calcium-dependent cell-cell adhesion molecule CDH1 150 (formerly known as E-cadherin or uvomorulin) [90,93–95]. Removing extracellular calcium 151 prevents CDH1 binding and causes embryos to de-compact [96–98]. The compaction process 152 was long thought to be driven by increased adhesion of cells via modifications of the CDH1-153 dependent adhesion machinery [1,99]. However, mechanical measurements in the mouse 154 embryo revealed that the forces driving compaction are located at the embryo surface rather 155 than at cell-cell contacts [11]. Micropipette aspiration of mouse blastomeres throughout 156 compaction uncovers raising tensions at the surface of the embryo as a result from the action 157 of the actomyosin cortex (Toolbox 1). Cells literally pull themselves together using their 158 intracellular muscles. The adhesion molecule CDH1 enables compaction by anchoring the 159 actin cytoskeleton of contacting cells so that they can effectively pull onto each other [67]. 160 CDH1 also provides local signals lowering contractility at cell-cell contacts and effectively 161 relaxing them [100]. Whether such mechanism is also responsible for human compaction 162 remains to be tested. Identifying the nature of the forces driving human embryo compaction 163 would help understanding what causes this process to fail in some instances. 164 Clinical studies found that compaction can be defective in several different ways (Fig 165 3). Compaction can be globally reduced with all blastomeres simply failing to grow their cell-166 cell contacts, which is associated with fewer embryos reaching blastocyst stage and lower 167

implantation rates [38,101]. Several cellular processes could be at fault, such as cell adhesion 168 or contractility. Micropipette aspiration on embryos failing to compact would narrow down the 169 options. Low tensions at the embryo surface would point towards contractility problems, such 170 as in mouse mutants for myosin paralogs [16,102]. High tensions at the embryo surface would 171 instead suggest defective adhesion, such as in mouse mutants for cadherin adhesion 172 molecules [11]. 173

Alternatively, compaction can be delayed. While a minority of embryo start compaction 174 as early as during the 8-cell stage, the majority of human embryos begin compaction during 175 the 4th wave of cleavages, with at least 9 blastomeres and most finish compaction during the 176 16-cell stage [10]. Embryos compacting less than 80 h after fertilization show higher rate of 177 implantation and live birth than embryos compacting after the 80 h mark [12,101,103]. What 178 controls the timing of compaction remains unclear though. In the mouse, activation of PKC can 179 induce a premature (and transient) compaction [104,105], which has been proposed to 180 promote contractility at the embryo surface [106]. 181

Finally, compaction can be unequal (also referred to as partial compaction), with some 182 blastomeres forming a compact mass while other cells appear excluded from this group (Fig 183 3). Excluded cells may later on contribute to the TE or be excluded entirely from the blastocyst. 184 The time of exclusion can occur before or after compaction is complete, with higher survival 185 rates for embryos excluding cells during the compaction process [9]. Interestingly, excluded 186 cells tend to be aneuploid while the rest of the embryo seems euploid [72]. This indicates that 187 exclusion of cells during compaction could serve as a repair mechanism ensuring that 188 defective cells do not participate to the blastocyst or to the embryonic tissues. Indeed, cells of 189 the placenta show a high rate of aneuploidy and have long been thought to act as a sink 190 eliminating defective cells from embryonic tissues [75]. However, the mechanism underlying 191 unequal compaction and exclusion of defective cells remain unknown. Again, excluded cells 192 could have lower tension at their surfaces or higher tensions at their cell-cell contacts, which, 193 if measured, would point at the underlying cellular process at fault, contractility and/or adhesion 194 (Toolbox 1). 195 196 Internalisation of the ICM precursors 197

The positioning of cells within the human embryo away from the cell-medium interface 198 is a critical morphogenetic step for the formation of the TE and ICM lineages (Fig 1). 199 Experimental manipulation of cell position reveals that outside cells from early human 200 blastocyst can adopt ICM fate if transplanted onto the ICM, away from the cell-medium 201 interface [18]. This apparent plasticity is explained by the fact that human lineages are 202 definitely set in the late blastocyst stage [20,31,107]. Until then, the position of cells within the 203 human embryo will guide their differentiation into either TE or ICM. How do blastomeres adopt 204 different positions within the human embryo in the first place? 205

Regardless of the molecular and cellular mechanisms that may control cell 206 internalisation, with sufficient cleavages, blastomeres could end up on the inside of the embryo 207 for geometrical reasons only [108]. Indeed, the packing of spherical objects into a sphere would 208 necessarily position cells on the inside when they become sufficiently small as compared to 209 the size of the embryo. If human embryos would be relying on geometrical packing only, then 210 reducing the embryo size would prevent the formation of an ICM. Dissociating human embryos 211 at the 4-cell stage reveals that 4-cell stage blastomeres are able to form smaller blastocyst 212 [21]. Quarter embryos compact, grow their lumen and contain inner cells expressing the ICM 213

marker NANOG. This implies that mechanisms other than geometrical packing drive cell 214 internalisation in human embryos. 215

In the mouse, at least two distinct mechanisms can internalise cells: oriented cell 216 divisions and contractility-meditated cell sorting. Oriented cell division consists in aligning the 217 mitotic spindle perpendicularly to the embryo surface so that one daughter cell is pushed 218 inward after cytokinesis [15,109,110]. Cell sorting occurs during the interphase and relies on 219 differences in surface tensions between cells, which, if sufficiently high, allow the strongest 220 cells to pull their way inside the embryo [16,17]. Both positioning mechanisms rely on the 221 establishment of apicobasal polarity with the apical domain facing the cell-medium interface 222 [13–16]. Division orientation is guided by the tethering of one of the spindle pole to the apical 223 domain [15]. This also facilitates the asymmetric inheritance of the apical domain, including 224 sub-apical components such as intermediate filaments when present [111]. The asymmetric 225 division of apical components, such as the apical kinases PRKCz and PRKCi (also known as 226 aPKCι and aPKCζ) that downregulate actomyosin contractility, among sister cells is key to 227 generate cell populations with different mechanical properties and drive cell sorting (Toolbox 228 1) [16,112]. Last but not least, apicobasal polarity governs the positional signals guiding the 229 differentiation of TE and ICM lineages (Toolbox 2). Apical signals promote TE lineages by 230 enforcing the nuclear localisation of the co-transcriptional activator YAP while cell-cell contacts 231 favour its cytoplasmic degradation [112,113]. Therefore, the apical domain is a master 232 regulator of lineage positioning and specification in the mouse embryo. 233

How much of this mechanism is conserved in human embryos is still unclear. Recently, 234 the presence and function of the core components of the apical machinery controlling TE fate 235 were confirmed in human and cow embryos [20]. Apical markers such as PRKCz, PARD6B or 236 AMOT are observed at the apical domain of human embryos (Toolbox 2). The accumulation 237 of PARD6B also seems to occur about 90 h post fertilization [114]. The human embryo then 238 consists of at least 16 cells and compaction has completed [10,12,103]. This is later than in 239 the mouse, which compacts and polarises during the 8-cell stage [11,20]. Similarly to the 240 mouse [112], inhibition of PRKCz using a chemical inhibitor or TRIM-AWAY, which targets 241 proteins to the proteasome using antibodies [115], impacts lineage specification in human 242 embryos [20]. When apical signals are reduced, YAP nuclear localisation is compromised and 243 the level of the TE-specific transcription factor GATA3 in outer cells is reduced [20]. Therefore, 244 the TE specification module by apical signals seems conserved in human embryos. However, 245 whether human embryos also rely on apicobasal polarity to positions inner and outer cells 246 remains to be determined. 247

248 At that point, the human embryo forms what is called the morula (Fig 1). The morula is 249

compact, surface cells have an apical domain facing the outside medium, which initiate their 250 TE differentiation, while inner cells do not have apical material and start adopting an ICM fate. 251 Importantly, compaction is functionally independent from polarisation (and cell internalisation). 252 This is best illustrated with mouse mutant embryos in which the processes can be impaired 253 selectively. Lacking essential apical kinases, Prkcz;Prkci mutants fail to form normal apical 254 domains, which prevents the correct specification of TE and ICM lineages but show a normal, 255 or even enhanced, compaction [112]. On the other hand, Myh9 or Myh9;Myh10 mutants fail to 256 compact due to the absence of sufficient actomyosin contractility but polarise correctly and 257 form ICM and TE lineages in the correct proportions [102]. Therefore, despite their apparent 258 synchrony, compaction and polarisation (and cell internalisation) can occur independently from 259

one another in mouse mutants. This also seems to be the case for human embryos, which 260 appear to compact normally when apical polarisation is impaired using inhibitors or TRIM-261 AWAY [20]. Whether compaction is required for human embryo polarisation remains to be 262 tested. 263 264 Lumen formation 265 Apicobasal polarisation is also key for the formation of the blastocoel, the first 266 mammalian lumen, which is the last morphogenetic step shaping the blastocyst (Fig 1). Fluid 267 accumulates within the morula and inflates the embryo to almost 10 times its original volume 268 [39]. Fluid is pumped into the intercellular space through the polarised surface epithelium while 269 inner cells cluster into a bud against the epithelium. This bud forms the embryonic pole of the 270 blastocyst, where the human embryo will mediate the uterine implantation [28,29,33,34]. In the 271 expanded blastocyst, surface cells are now differentiated into TE and inner cells into ICM, 272 which further specifies into PrE and Epi as the blastocyst prepares for implantation. 273 The formation of the blastocoel relies on the tight sealing of the TE, polarised transport 274 and the ordered detachment of the ICM into one pole. Recent studies in the mouse provided 275 mechanistic details on how these architectural changes proceed. Initially forming a domain in 276 the centre of the cell-medium interface of surface cells, the apical domain expands [15]. When 277 the apical domain hits the apical edge of cell-cell junctions, tight junctions seal the TE [24]. 278 The expansion of the apical domain relies on coordinated cytoskeletal actions of the 279 microtubules signalling to exclude acto-myosin from the apical domain, as observed in PRKC 280 mutants [16,24]. This sealing can be challenged during cell divisions, which requires cells to 281 round up and pull on their neighbouring cells, putting junctions under mechanical stress 282 [22,116]. Therefore, tight junctions must reinforce themselves to prevent the embryo from 283 collapsing under the increasing pressure of the blastocoel and during the waves of divisions 284 of the TE. Polarised transport occurs through the cells rather than via junctions, as suggested 285 by mouse mutants that, after failing all successive divisions, form a single-celled embryo, which 286 nevertheless initiates blastocoel formation [102]. Indeed, as long as osmolytes transporters, 287 such as the Na/K pump, and aquaporins are polarised along the apicobasal axis, blastomeres 288 pump fluid from their apical to their basolateral compartment. This basolateral compartment is 289 enriched in adhesion molecules, such as CDH1, and mechanically opposes detachment and 290 fluid accumulation [23]. However, the fluid pressure, between 5-10 times higher than that of 291 cells, is large enough to fracture cell-cell contacts. This includes contacts between ICM cells, 292 which are transiently broken [23]. Contacts then repair themselves thanks to the action of their 293 actomyosin cytoskeleton [117]. Interestingly, patterning the contractility of cells is sufficient to 294 direct the positioning of the lumen and ICM, effectively dictating the first axis of symmetry of 295 the mammalian embryo, which guides its implantation. 296 How much of these mechanisms are conserved in human embryos is unclear. The 297 presence of key components from the tight junctions and fluid pumping machineries are likely 298 [19,118–121]. More generally, the differentiation of the TE into a functional epithelium is 299 required for the integrity of the human blastocoel, which fails when key transcription factor 300 POU5F1 is mutated [19]. If conserved in human embryos, these mechanisms could have 301 important clinical implications for better assessing embryos. The rate of blastocyst expansion 302 is thought to provide information on its implantation potential, with implanting embryos showing 303 higher expansion rates than non-implanting embryos [37,39]. Interestingly, the expansion rate 304 is also an indicator of the ploidy state of the embryo, with aneuploid embryos inflating slower 305

than euploid ones [122]. Aneuploidies seem to be more frequent in the TE directly lying over 306 the ICM, the polar TE, which attaches to the uterus [123]. The reason for the slower expansion 307 rate could be linked to leaky TE, for example as a result of poor sealing of tight junctions or 308 could be due to weaker polarised transport. Non-invasive methods allowing measuring the 309 sealing and transport of human embryos would be very helpful to assess these properties and 310 better understand the causes underlying slower expansion. Fluorescence based assays can 311 be used to detect leakage [22,68]. Measuring volume changes after osmotic shocks would 312 allow assessing the transport properties [124]. Besides, analysing blastocyst growth rates with 313 higher temporal resolution could give more information on sudden collapses [55,116]. Several 314 studies investigated the relationship between collapses and implantation rates, with 315 sometimes contradictory conclusions [125–127]. As collapses may result from cell divisions 316 transiently disrupting the TE seal, they may report on a healthy dividing TE [22,116]. 317 Alternatively, if collapses are too frequent, they may reflect a poorly sealed and therefore 318 dysfunctional TE. Finally, reports of human blastocyst with multiple clusters of inner cells 319 suggest that monozygotic twinning may originate from the blastocyst [128]. If conserved in 320 human embryos, hydraulic fracturing of contact between inner cells during blastocoel initiation 321 may be responsible for the formation of multiple ICM. However, the formation of multiple ICM 322 was not observed in the mouse, even when the fracturing or repair mechanisms were 323 genetically manipulated [23]. Further investigations in the adhesive properties of ICM and TE 324 cells will be needed to identify how the ICM could split. 325 326 Perspectives 327 Based on recent studies in mouse and human embryos, we have detailed how key 328 steps of the shaping of the human blastocyst could proceed and fail. Since morphology serves 329 as the basis for selecting the most suitable human embryos during assisted reproduction 330 technology procedures [35,36], we believe there is much to be learned from the mechanisms 331 of blastocyst morphogenesis. Current and future studies are taking exciting directions with the 332 development of artificial intelligence (AI) based image analysis and non-invasive mechanical 333 measurements. AI already shows performances equivalent to or even better than clinicians in 334 predicting implantation rates, based on retrospective studies [40–42]. Most importantly, AI will 335 allow more systematic procedures between clinics, as embryo diagnostic will rely less on the 336 subjectivity of clinician eyes. However, algorithms, especially those based on machine 337 learning, do not necessarily permit knowing the precise criteria used for predicting the 338 implantation potential of embryos. Therefore, we should not rely on AI to uncover mechanistic 339 understanding underlying developmental defects. On the other hand, non-invasive mechanical 340 measurements offer the possibility to better understand the mechanisms for a given 341 morphogenetic defects. For example, micropipette aspiration has been used on mouse and 342 human oocytes and provided improved prediction of developmental potential [129]. 343 Micropipette aspiration was performed through the zona pellucida, a porous glycoprotein shell 344 of high elasticity compared to the oocyte. This aspiration measurement is therefore likely to 345 reflect how porous and elastic the zona pellucida is, rather than probing the mechanics of the 346 oocyte. Nevertheless, similar measurements could reveal how compaction fails in human 347 embryos or even specific blastomeres. Such microaspiration could for example be performed 348 as blastomeres are being remove during preimplantation genetic diagnostics. Together, better 349 understanding the shaping of human blastocyst will improve the success rates of ART 350

procedures, which become more frequent as the age of first conception keeps on increasing 351 [130]. 352

Acknowledgements 353 We thank Diane Pelzer and Markus Schliffka for critical reading of the manuscript. J.F. is 354 funded by the Fondation pour la Recherche Médicale. Research in the lab of J.-L.M. is 355 supported by the Institut Curie, the Centre National de la Recherche Scientifique (CNRS), the 356 Institut National de la Santé Et de la Recherche Médicale (INSERM), and is funded by grants 357 from the ATIP-Avenir program, the Fondation Schlumberger pour l’Éducation et la Recherche, 358 the European Research Council Starting Grant ERC-2017-StG 757557, the European 359 Molecular Biology Organization Young Investigator program (EMBO YIP), the INSERM 360 transversal program Human Development Cell Atlas (HuDeCA), Paris Sciences Lettres (PSL) 361 “nouvelle equipe” and QLife (17-CONV-0005) grants and Labex DEEP (ANR-11-LABX-0044) 362 which are part of the IDEX PSL (ANR-10-IDEX-0001-02). 363

Figure Legends 364

365 366 Figure 1: Human blastocyst morphogenesis 367 Starting from the zygote (Day 0), the human embryo undergoes cleavage divisions, which 368 become progressively less synchronous with successive waves of cleavages, finishing here 369 with the 6th wave on Day 5. During the cleavage stages, there is no morphogenesis. 370 Compaction begins on Day 3, around the time of the 4th wave and ends on Day 4. The morula 371 is complete when the embryo is compacted and contains inner cells (red). At the time of 5th 372 cleavage, the outer cells (blue) pump fluid to inflate a lumen and turn the embryo into the 373 blastocyst. Defective morphologies result from defects in cleavages (see Fig 2), compaction 374 (see Fig 3) and/or lumen formation. Note that developmental time is not represented linearly. 375

Day 0 Day 1 Day 2 Day 3 Day 4 Day 5

Lumen formation

Cleavage defects Compaction defects Lumen defects

Cleavage stages Morula Blastocyst2-cell 4-cell 8-cell

Cleavage2nd 4th 5th 6th1st 3rd

Compaction

Internalisation

16-cell 32-cell

376 377 Figure 2: Cleavage defects during human preimplantation development 378 Normal cleavage division splits the volume of the blastomere in equal parts (top). Cleavage 379 defects can affect the volume of daughter cells in a noticeable way (right). Cytokinesis itself 380 can be asymmetric and produce daughter cells of unequal size. Tripolar mitotic spindles can 381 produce three daughter cells instead of two and result in what is referred to as direct cleavage. 382 This often result in the production of aneuploid cells (marked by the presence of extra 383 chromosomes in red). Cytokinesis or abscission can fail resulting in the fusion of sister cells 384 and the formation of a polyploid or bi-nucleated cell (reverse cleavage). During cytokinesis, 385 cellular fragment frequently form. Fragments waste material and sometimes entrap 386 chromosomes leading to aneuploidy. 387

Defective cleavagesNormal cleavage

Unevencleavage

Direct cleavage

Reverse cleavage

Fragmentation

388 389 Figure 3: Compaction defects in human embryos 390 During normal compaction (top), the human embryo increases the area of contact between 391 cells and reduces the surface exposed to the outside medium to form the compacted morula. 392 Compaction can fail completely or be weak (bottom left). Compaction can also occur in subset 393 of blastomeres only, excluding some blastomeres (bottom right). Blastomeres excluded from 394 unequally compacted morula can be aneuploid. Excluded blastomeres could contribute to the 395 TE and therefore placental tissues or be excluded from the blastocyst entirely. 396

8-cell Compaction initiation Compacted morula

No compaction Unequal compaction

Compaction

Defectivecompaction

397 398 Toolbox 1: Surface tensions during compaction and cell internalisation 399 Comparing the human embryo to contacting soap bubbles provides a general theoretical 400 framework explaining compaction and cell internalisation [11,16,108]. Blastomeres are given 401 a surface tension at their cell-medium (γcm) and cell-cell interfaces (γcc). Following the Young-402 Dupré equation, the level of compaction is simply given by the ratio α of tension between cell-403 cell and cell-medium interfaces (γcc/2γcm). The surface tension of blastomeres can be 404 calculated using the Young-Laplace equation, which relates the surface tension, pressure and 405 curvature of liquid-like materials. Micropipette aspiration can then be used to determine the 406 pressure inside blastomeres [11,131]. This approach revealed that, during compaction of the 407 mouse embryo, γcm doubles while γcm decreases by a third. Using the absolute values of 408 surface tensions further uncovers that ¾ of mouse compaction results from the changes in 409 surface tension γcm and ¼ from the relaxation of cell-cell contacts [11]. 410 Considering the ratio of surface tensions at cell-medium interfaces of individual blastomeres δ 411 = γcm1/γcm2, internalisation occurs when cell 1 grows its tension above a threshold value. 412 This threshold is set at δ > 1 + 2α, and therefore depends the level of compaction [16]. 413 Interestingly, differences in size have no effect on internalisation in theory. For the mouse 414 embryo, the internalisation threshold is at 1.5, i.e. cells need to grow their tension 50% higher 415 than that of their neighboring cells to pull their way inside the embryo. 416

Compacted

Compaction parameter ! = "cc /2"cm

"cm2"cm1

"cc

Internalisation

Internalisation parameter# = "cm1 /"cm2

"cm "cm

"cc

Not compacted

↑ "cm

↓ "cc

417 418 Toolbox 2: De novo apicobasal polarisation and TE-ICM specification 419 During preimplantation development, blastomeres form de novo a domain of apical material 420 (red) at the surface of the embryo, away from cell-cell contacts (basolateral interfaces in blue). 421 The apical domain forms in the center of the contact free surface and signals to the actomyosin 422 cortex (green) to deplete it locally. The apical domain progressively expands until it reaches 423 the apical edge of the cell junctions. The apical domain contains molecules such as CDC42, 424 PARD6B or PRKC that prevent the LATS kinase from phosphorylating the co-transcriptional 425 activator YAP. YAP can shuttle to the nucleus to interact with the transcription factor TEAD4 426 and activate the expression of TE specific genes such as GATA3. Without an apical domain, 427 inner cells show less YAP in their nucleus as the LATS kinase, with the help of NF2 and AMOT 428 localised at cell-cell contacts, can phosphorylate YAP and target it to the proteasome. The 429 presence of some of the signalling molecules involved in the subcellular localisation of YAP 430 has been confirmed in human embryos (labelled in bold font), while other have only been 431 studied in mouse embryos. 432

Surface

Inside

Apical

Basolateral

AMOT

LATSAMOTNF2 P

TEAD4

TEAD4

YAPP

YAP

YAP

CTNNB

CTNNB

LATS

CTNNB

CTNNB

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